This invention relates to a crustacean larva raising method and apparatus.
This invention has particular but not exclusive application to larva raising method and apparatus for use with Thenus spp., and for illustrative purpose reference will be made to such application. However, it is to be understood that this invention and inventive elements thereof could be used in other applications, such as rock lobster and slipper lobster larvae.
There have been many attempts made to develop larva-rearing strategies for commercial species of crustacea. To date these have been concentrated on developing strategies for species of rock and slipper lobster larvae. A summary of these processes is given in Table 1.
A chronology of larval efforts is given in Table 2.
The present four major commercial-research rearing tank systems for rock and slipper lobster larvae are, for the Southern rock lobster, (Jasus edwardsii) the systems developed by the Tasmanian Aquaculture and Fisheries Institute (TAFI), for J. edwardsii and the Eastern rock lobster, (Jasus verreauxi) the systems developed by the National Institute of Water and Atmospheric Research (NIWA) of New Zealand, for the Japanese rock lobster, (Panulirus japonicus) the system developed by Fisheries Research Institute of Mie (FRIM), the Japan Sea-Farming Association (JSFA) and Research Institute for Science and Technology of The Science University of Tokyo, all of Japan. The broad features of these systems and the results published are as follows:
1. (TAFI)
Southern rock lobster
10 l water in 30 l tanks
Stocking density of 20 newly hatched larvae per l
No juveniles obtained.
2. Upwelling tank system (NIWA)
Southern rock lobster
Combination of 4xc3x9772 l tanks
Stocking density of 26 newly hatched larvae per l
Only one juvenile survived in 1990
3. Zero water movement tank system (FRIM)
Japanese rock lobster
150-180 l in 200 l tanks
Stocking density of 20 newly hatched larvae per l
Less than 1% survival to the juvenile stage (up to 10 juveniles)
4. (JSFA)
Japanese rock lobster
150-180 l in 200 l tanks
Stocking density of 20 newly hatched larvae per l to 1 final stage larvae per l
Approximately 1% survival to the juvenile stage (up to 100 juveniles)
Thenus spp., commonly known as Moreton Bay bugs, Slipper lobsters and Bay lobsters, are found along the entire northern coast of Australia from Shark Bay in Western Australia to Coffs Harbour in northern New South Wales (Kailola et al., 1993). There are two Thenus species: Mud bugs (Thenus sp.) and Sand bugs (Thenus orientalis). Mud bugs are brown overall and have brown stripes on their walking legs, while Sand bugs are speckled overall and have spots on their walking legs. Mud bugs prefer a bottom of fine mud, and are typically trawled from inshore coastal waters of 10 to 30 meters depth. Sand bugs tend to prefer sediments with a larger, coarser particle size, and are usually trawled from a depth of 30 to 60 meters in the coastal shelf and offshore areas.
Currently, commercial aquaculture of Moreton Bay bugs is not being carried out anywhere in the world. The major hurdle in commercialisation is the difficulty in maintaining the bugs through the larval stages. Like other slipper or rock lobster species, Moreton Bay bugs have a very characteristic flattened larval stage called the phyllosoma. They circulate in the plankton, rising and falling in the water column, and this makes it difficult to adjust the culture environment.
Recently however, a comprehensive study of the culture conditions of Moreton Bay bug phyllosomas was undertaken, suggesting high potential for commercial aquaculture of these species (Mikami, 1995). Phyllosomas of Moreton Bay bugs pass through four larval stages in 25 to 35 days, with a high level of survival on a small scale and take one year to achieve commercial size (250 g).
Following the study by Mikami (1995), further research has been undertaken by the present applicant over the past five years. The major aim of this study is the commercialisation of Moreton Bay bug aquaculture from the small, experimental scale. To date, the single most important issue has been to solve the technical aspects of Moreton Bay bug larval rearing.
In one aspect this invention resides in a crustacean larva raising method including the steps of:
providing a tank adapted to hold larva raising medium to a depth of at least 10 cm;
continuously supplying substantially sterilized, filtered larva raising medium to said tank through a plurality of outlets disposed about the tank and adapted to cause horizontal circulation of said medium and having an outlet flow velocity selected to prevent larva damage;
continuously draining said medium through a drain assembly including a larva screen having a flow velocity of said medium therethrough selected to prevent damage to larvae, and
maintaining said medium at a temperature selected to accommodate the larva species to be raised.
In a further aspect, this invention resides broadly in crustacean larva raising apparatus including:
a supply of substantially sterilized, filtered larva raising medium;
a tank adapted to hold said larva raising medium to a depth of at least 10 cm;
a plurality of outlets connected to said supply and adapted to deliver and cause horizontal circulation of said medium in said tank;
drain means having a larva screen and configured to maintain a selected level in said tank, and
temperature control means for said medium.
The larval rearing tank may be round or oval in horizontal cross section such that a continuous one-way circulation may be maintained. Alternatively, the larval rearing tank may comprise an annular tank. As a yet further alternative, the larval rearing tank may comprise an annular raceway having straight portions closed by end portions. Preferably, the raceway comprises a modular construction of curved and straight portions, whereby the linear dimensions and thus holding capacity may be selected. For example, the modular components may be moulded in plastics material and be adapted to be bolted up in assembly to form the raceway. The modular components may be provided with preformed joint sealing, or may in the alternative be sealed with an in situ cast sealing such as curable silicon or other sealant.
The tank depth is preferably less than one meter. Preferably, the water depth is maintained at about 10 to 20 cm. This relatively shallow depth will allow increasing feeding frequency of larvae. In the case of the preferred annular and or modular raceway construction, the section of the raceway may be for example 30 cm deep. Whilst the width of the section may be of any suitable dimension determined at least in part by the arrangement of the rearing media outlets, it is preferred that this dimension also be in the region of 30 cm.
For phyllosomas of Moreton bay bugs, a typical stocking density of about 40 newly hatched larvae per l is used, gradually reduced to about 10-15 final stage larvae per l.
In order to increase floor area densities in industrial situations the tanks may be arrayed in stacks.
The medium will be selected according to the species to be raised. In general the medium will be seawater or synthetic seawater of composition selected to match the natural environmental medium in which the organism exists in the wild.
The water outlets may comprise a plurality of nozzles. The nozzles may be of a number selected to encourage the continuous one way circulation with consistent flow about the circuit. The number of nozzles and capacity of tank can be used to adjust the volumetric flow rate.
The flow velocity of the nozzles may be any flow velocity selected to maintain circulation in the tank whilst avoiding shearing injury to the larvae. The flow velocity is preferably maintained in the region of below 4 to 6 m per minute at least for early larval stages such as the at the 1 st phyllosoma stage of Moreton bay bugs. Preferably the flow rate is the minimum flow rate consistent with maintaining circulation of the medium in the tank.
The outlets may be located at the bottom of the tank or the top of the tank. For example, the outlets may be associated with a linear or ring mains manifold located at the bottom of the tank or at any position up the walls of the tank including above the medium level. There may be provided a single manifold or a plurality of manifolds. In one embodiment of the invention associated with the preferred modular raceway tank, the outlets are tubes with 4 mm nozzles extended from 19 mm polyethylene manifolds disposed on the upper portion of the inner and outer walls of the raceway and installed after assembly of the modular structure. The outlets extend down to the bottom of the tank and the nozzles are aimed in the direction of desired circulation and preferably directed somewhat inwardly away from the walls.
The medium may be supplied by a continuous one way system or may utilize some recirculation. For example, in a one way supply, the medium such as seawater may be filtered from a natural supply through a 1 xcexcm filter and preferably a 0.5 xcexcm filter in a header tank.
The drain means is preferably provided with a mesh size of about 1 mm. The flow rate across the mesh may be determined in each case empirically. However, it is preferred that the flow rate per unit area be much less than the inflow velocity and accordingly the surface are of the mesh is preferably maximized. The drain means may be used to maintain the level of medium in the tank. To this end the drain means may comprise a surface drain set to the desired level or 10-20 cm. Alternatively, the drain means may be located at any level in the medium column, whereupon the level may be medium supply/drain rate controlled. In cases where the drain controls the level, this may be provided by means of a meshed drain inlet of enlarged area relative to a standpipe taking the medium to waste or recirculation, which standpipe may be adjustable in length.
The tank may be provided with a cover or other means selected to occlude light from the tank at selected intervals. Light should be excluded from the tank in the daylight hours to maximise feeding frequency of the larvae. To this end it is preferred that the tank be formed of an opaque material.
Contamination by bacteria, protozoa or fungi is a serious problem for phyllosoma rearing. The major sources of contamination for larval rearing are incoming water, food, the air, human handling and the starter culture (eggs, water and newly hatched larvae from the hatching tank). Larval rearing water should be kept free from any organisms. Accordingly, after filtration the filtered seawater may then be sterilized by any suitable means such as UV sterilization, submicron filtration, chlorination, acidification or ozonisation. For example, the filtered seawater may be exposed to UV radiation from an arc or other source at above about 1 l/hour/Watt level to minimise bacteria.
Alternatively, chlorine at about 10 ppm may be maintained in incubation with the medium for about 12 hours, preferably without aeration, followed by the addition of sufficient sodium thiosulfate to neutralise the chlorine.
The temperature of the filtered, sterilized supply may be maintained in the desired range by any suitable means such as heaters and/or chillers with appropriate thermostats.
In the alternative to the one way system, there may be provided a semirecirculation system including the larval rearing tank and two or more sub tanks. Preferably, two sub tanks are used. The sub tanks are each of at least the same capacity as the larval rearing tank. In this embodiment, the sub-tank containing filtered, sterilized water may be circulated into the filled larval rearing tank for about 24 hours residence using a submersible pump. After 24 hours, the pump may be transferred to the other sub-tank, with the water controlled at about the same temperature, preferably within about xc2x10.5xc2x0 C. Water may then be circulated into the larval rearing tank again. Preferably, the flow rate is the same as the one way flow-through system. While water in one sub-tank is being used, the other sub-tank may be emptied and dried.
In the case of semirecirculation, the rearing water may be sterilized by, for example, 10% chlorine for a period of 12 hours followed by neutralisation with 10% sodium thiosulphate. Preferably, the rearing water is tested such as by Palintest(copyright) (DPD No 1) before introducing into the rearing system to make sure no chlorine remains.
In the case of phyllosomas of Moreton bay bugs, the temperature range of the medium is preferably between 26-27xc2x0 C. Phyllosomas can be reared at temperature ranges between 24-30xc2x0 C., but temperatures lower than 26xc2x0 C. will result in a slower growth rate, and those higher than 28xc2x0 C. will increase the risk of unsuccessful moulting, cannibalism and disease. When larvae are transferred to the rearing tank, it is preferable to keep the temperature of the larval rearing system at substantially the same temperature xc2x10.5xc2x0 C. as the source of the larva such as a hatching tank. If rearing water temperature has to be changed, temperature variation is preferably kept within 1 degree per hour.
The salinity of the medium may vary according to species. In the case of phyllosomas of Moreton bay bugs, the salinity may be maintained in the range of between 25-40 ppt and preferably between 34-36 ppt. Phyllosomas are extremely intolerant to sudden changes in salinity, so salinity change should be kept within xc2x11 ppt per day.
Throughout all phyllosoma stages, phyllosomas show strong photopositive reaction. To avoid congregation of larvae at the surface during the daytime, the rearing system may be covered such as by a black plastic sheet.
The level of pH may be kept at between 7-9, and preferably between 8.2-8.5 being the natural seawater pH level.
Strong aeration damages larvae, so it is preferred to avoid using aeration in the rearing tank. The oxygen level of the rearing water is preferably kept at more than 7 ppm, at 26-27xc2x0 C. Larval oxygen consumption is very low, so the circulation of rearing water with a large surface area is generally adequate for supplying larval oxygen demand without aeration, with control of stocking density.
Under the flow-through system, the preferred maximum rearing densities of phyllosomas are:
40 first instar larvae per l
25 second instar larvae per l
10 third instar larvae per l and
5 fourth instar larvae per l.
Rearing density levels higher than this may result in a high level of cannibalism at the time of moult. Pre-moult/post-moult larvae are eaten by intermoult larvae.
The phyllosomas are preferably fed a controlled diet. The maximum phyllosoma growth and survival has been determined to be obtained by the use of chopped, fresh, live mollusc flesh, preferably live pipis (Donax spp.). Using frozen food will result in a slower growth rate than fresh food. Brine shrimp (Atemia spp.) can also be used, but only for 1st instar phyllosomas.
The use of live pipis occasionally causes a high level of mortalities at the time of moulting. This is called moult-death syndrome (MDS). The cause of MDS amongst other species is still unclear, but in the case of Thenus, MDS is related to seasonal variances in food quality. To obtain a standard quality of food, enrichment of bivalves is preferred.
Use the green micro-algae Nannochloropsis spp. or other micro-algae and/or diatom species such as Isochrsis spp., Chaetoceros spp. and Pavlova spp. for enrichment has proven useful. Enrichment may comprise culturing pipis at 25-28xc2x0 C. with algae water at a cell density of preferably greater than 20xc3x97107. For example, there may be used approximately 1 kg of pipis (wet weight with shell) per 40 l of algae water. Preferably, the water is replaced every 12 hours. The enrichment process may be conducted for at least 24 hours and preferably 48 hours. The level of ammonia in the algae water should be maintained below 1 ppm. Flesh content of pipis (gut, gonad, gill and mantle) is approximately 20% of total weight.
Alternatively to the preferred algae, dried commercial species may be used such as Marine Sigma (Nisshin Science), Marine Growth (Nisshin Science), and Algamac-2000 (Bio Marine). The number of cells of these commercial products should be kept at  greater than 20 million per ml.
Food preparation may be by any suitable means. In the case of the preferred pipis, the flesh may be chopped roughly followed by washing through at least two grades of mesh, such as 0.5 to 2.0 mm for a first wash and thereafter  less than 0.5 mm. The large mesh size is preferably varied according to larval stage. For example, there may be used 1.0 mm for the first instar, 1.5 mm for the 2nd instar and 2.0 mm for the 3rd and 4th instars. The pieces of chopped flesh retained between the large and small mesh sizes may be set aside. The pieces of flesh retained in the large size mesh may be chopped again, repeating the above process.
Food must be sterilised before feeding in order to avoid bacterial contamination. For example, the flesh may be washed in UV sterilised seawater carefully, and then incubated in 0.1% chlorine seawater solution for a period of 30 minutes or more. Then wash the food particles by UV sterilised seawater again on the small mesh before feeding to larvae.
Prepared food materials with seawater may be distributed equally in the rearing water using for example a pipette. Food particles will sink to the bottom of the rearing tank. Food particles remaining in the rearing tank after feeding should be cleaned out before adding the next lot of food. The feeding level changes depending on growth stage and intermoult stage. The level of feeding should be adjusted by taking note of how much food remains from the previous feed.
Phyllosomas start eating from the night of hatching. To obtain synchrony of larval moult, it is preferred to not feed on the morning of Day 1. As the phyllosomas start eating more, so preferably adjust the feeding level depending on level of remaining food. Feed preferably twice a day in the early morning and late evening. At Day 5-6, the phyllosomas start preparing to moult, so the feeding level may be decreased from the evening of Day 5.
First instar phyllosomas moult to the second instar in the early morning, so the feeding level in the morning may be minimised, with more in the late evening. On days 7-9, feeding twice a day is still acceptable but towards Day 9 phyllosomas start to eat more. Monitoring the level of remaining food regularly is preferred to avoid starving the phyllosomas, feeding 3 times a day if necessary. On days 9-10, feeding levels will still be high, even before moulting. It is preferred to make sure enough food is available through the nights to avoid cannibalism in the morning.
Larvae usually moult to the third instar in the early morning (4-5 am), and therefore it is preferred to make sure enough food is available before and during the moulting stage. An extra feeding before moulting is desirable, if there is no food remaining in the tank. Post-moult stage larvae will not eat food for 3-6 hours, and therefore the morning feed should be minimised, with a higher level in the afternoon. From Day 12-16, larvae may be fed 3 times a day, preferably every 8 hours such that food is always available. Starvation of phyllosomas will cause a high level of cannibalism when third instar phyllosomas moult to fourth instar phyllosomas.
Fourth instar phyllosomas (pay 15-27) may be fed over Days 15-17 at three times a day, preferably making sure food is always available. At days 18-21, the feeding level of phyllosomas is now at its peak. Larvae may be fed three times a day or more, preferably without delaying any of the three feedings for more than 2 hours. From day 21-30, phyllosomas start to metamorphose to the nisto stage, and therefore the feeding level should be decreased with the decreasing number of fourth instar phyllosomas. When phyllosomas are not eating food around day 25-26, feeding can be reduced to only twice a day.
Under optimal rearing conditions (physical and nutritional), intermoult periods of phyllosomas are preferably synchronised. The timing of these moultings depends on the rearing conditions (temperature, food condition, stock density and so on), and therefore it is preferable to maintain optimal rearing conditions throughout all larval stages.
Larvae can be reared using only one rearing tank in accordance with the present invention, with no tank exchange required. When fourth instar phyllosomas metamorphose to the nisto, pre-metamorphosis phyllosomas should be transferred to the nisto tank. Metamorphosis always occurs in the late evening just after sunset. Pre-metamorphosis phyllosomas can be identified by changes in the external morphology: the appearance of xe2x80x9cWxe2x80x9d shaped gaps at the basement of the antenna (these become the eye sockets); small dots on the carapace; and the changing of body colour to white. Pre-metamorphosis phyllosomas should be transferred with seawater to the nisto tank.
To avoid bacterial contamination, human contact with larval rearing water must be avoided. For example, it is preferred to wash hands with an anti-bacterial soap before the treatment for larvae. Plastic instruments may be kept, for example, in a 0.01% chlorine water bath when they are not being used. Preferably, change the water completely every 3-4 days. Glass instruments may be washed carefully with fresh water and keep on a shelf when dry.
Phyllosomas can start eating immediately after hatching, but this depends on yolk retention of larvae and temperature. In general, phyllosomas start eating 6-12 hours after hatching, but can survive for up to 72 hours without food. Starvation time of up to 48 hours at 27xc2x0 C. will have no influence on survival and moulting. The 50% level of Point of No Return (PNR50) is generally 72 hours after hatching, but this will change depending on yolk retention of larvae. Delaying the initial feeding will prolong the duration of the first instar. After moulting to the second instar, the initial starvation will have no further influence on growth.
Phyllosomas eat less food at pre- and post-moulting stages (xc2x112 hours of moulting) than during the middle of the intermoult periods. In the middle of the intermoult periods, phyllosomas eat constantly, day and night. Although phyllosomas have a strong capacity for starvation and can survive for more than 72 hours without food, long-term starvation and lower feeding levels will result in an increased risk of MDS at the time of moulting.
Phyllosomas are not passive feeders; they approach and attack prey. The phyllosomas attack (pick up) the food using pereiopods, and pass to the mouthparts, located at the central part of the carapace (ventral side). The mouthparts comprise the labrum, pared paragnaths, mandibles and 1st maxilliped. The labrum and paragnaths cover the top of the mandibles. The food particles are pushed onto the paragnaths by the 1st maxilliped, and cut roughly into small food masses. Then the mandibles, which have a scissor-like structure on the anterior-tip, break down the food into even smaller pieces. Therefore phyllosomas can only eat soft food masses with a high water content.
After phyllosomas ingest food materials into the gut system, the colour of the midgut gland changes from transparent to white, due to the appearance of lipid rich globules within the cytoplasm of the midgut gland cells. Only a portion of the ingested food materials goes into the midgut gland area, where the main part of digestion occurs. The majority of food materials pass through the midgut tubule and are excluded via the anus, 5 to 10 minutes after eating. Faeces of phyllosomas are lipid-rich pseudo-faeces.
Phyllosomas are plankton, and usually swim in the same direction as the water current. However, phyllosomas are also strongly photopositive and can swim across a water current of 10-15 m per minute towards a light source. In the hatchery, phyllosomas congregate at the spot where light intensity is the highest during the daytime, but spread themselves evenly in the water column during the night-time. Phyllosomas show a strong photopositive phototaxis even under illuminance levels of 0.5 xcexcEmxe2x88x922secxe2x88x921. Phyllosomas can also swim to the bottom of the tank and pick up food materials against a water current of 10-15 m per minute. When phyllosomas are healthy, they swim with rotation of their bodies.
Moulting usually only occurs in the early morning around sunrise. The pre-moult stage (where the internal chemical composition of phyllosomas change) starts 2-3 hours before the actual moulting. The pre-moult stage larvae can be identified by a change in body colour (transparent to white-pink) and swelling of the carapace. Post-moult phyllosomas are very soft and fragile for 2-3 hours. Movements of post-moult stage phyllosomas will depend on the water current. Cannibalism occurs only at the time of moulting, when intermoult larvae eat post-moult and pre-moult stage phyllosomas. Post-moult phyllosomas start to eat food 2-4 hours after moulting.
Metamorphosis only occurs in the late evening around sunset, with the process lasting only 10-20 minutes. Pre-metamorphosis phyllosomas can be distinguished by their external morphology: small dots on the middle of the carapace (these become the top edge of carapace after metamorphosis) and xe2x80x9cWxe2x80x9d shape gaps at the basement of antenna (these become the eye sockets). The entire body of pre-metamorphosis phyllosomas is thick and tends to be bright-white.
Cannibalism of phyllosomas can be observed only at the time of moulting, where intermoult stage phyllosomas eat pre- and post-moult stage phyllosomas. However, if nutritional requirements of phyllosomas are satisfactory throughout the intermoult period, the level of cannibalism should be minimised.
The causes of phyllosoma diseases can be divided into bacterial, fungal, nutritional, viral and environmental or stress origins. Contamination by bacteria is the major problem in rearing phyllosomas. The most common sources of bacterial contaminations are starter culture (eggs), incoming water and foods. Sterilisation of the seawater by physical methods, such as filtration, UV, and/or chemical methods, such as chlorination and ozonisation, is effective in preventing diseases of bacterial origin. There are a number of critical diseases originating from bacteria.
Toward the end of larval rearing (amongst fourth instar phyllosomas), Vibrio infection will result in gut blockage. The symptom is accumulation of food material in the midgut tubule (constipation). Larvae will die after 6-12 hours. The midgut path is infected by Vibrio, and gut material is no longer excluded from the midgut tubule. This disease is not contagious. An accumulation of bacteria on the bottom surface of the rearing tank is thought to cause this disease. Daily maintenance, particularly of the bottom surface, is an important factor in prevention. Alternatively, exchanging the rearing tank can reduce mortalities from this disease.
The pared antenna glands are an excretory organ located at the base of the antenna. The antenna glands are surrounded by a single cell layer called a bladder, where ammonia is selectively transported from the hemolymph. The bladder is connected to the opening at the surface of the carapace, and ammonia is excreted via the opening. The diameter of this opening is less than 5 xcexcm. Because of a high level of ammonia around the opening of antenna glands, filamentous bacteria can grow easily and shade the opening of the antenna gland, causing necrosis at a cell layer. Healthy antenna glands are transparent, but their colour will change to brown/black after necrosis. If both antenna glands are infected, larvae will die after 24-48 hours. Streptomycin at 10 ppm can prevent bacterial growth, but there is no cure for necrosis of the antenna gland. Daily cleaning of the larval tank system is an important means of reducing mortalities caused by this type of disease.
Filamentous (Leucothrix sp) bacterial infection can be observed on the surface of the exoskeleton. Poor water quality management is the cause. Though streptomycin sulphate at 10 ppm is an effective preventative measure, the continual use of antibiotics should be avoided.
Fungal origin diseases may include infections from marine fungi commonly found in seawater, and occasionally grow on the surface of the larval exoskeleton, particularly on the exopodal setae of periopods. Marine fungi start to grow 3-4 days after hatching and moulting. Fungi growing on the exopodal setae sticks will attract food particles, and as a result larvae will be xe2x80x9cgluedxe2x80x9d together. These larvae will not die immediately, but feeding and swimming will be destructive, causing high mortality two or three days after the appearance of the fungi. Formalin at 20 ppm twice a day can prevent marine fungi growth, but will not be effective against fungi already on the surface of larval exoskeleton.
Nutrition is an important factor for not only obtaining optimal growth, but also for prevention of any kind of disease. Phyllosomas should have some degree of resistance against diseases of bacterial origin if their nutritional requirements have been adequately met. Moult Death Syndrome (MDS) is a catastrophic syndrome, observed at the time of moulting. Larvae simply stop moulting in the middle of the process and die. This syndrome cannot be predicted until the time of moulting. During intermoult periods, larval survival and activity are always high, and intermoult periods are usually synchronised. Seasonal variation in the nutritional content of natural foods (bivalves, pipis) is considered to be the major cause of MDS. During early spring (September) to mid-summer (December), a high level of MDS is observed with the feeding of non-enriched pipis. MDS levels are much lower between mid-summer (January) and autumn (June).
Enrichment of pipis (improvement of nutritional value) as described above is an effective way to prevent this syndrome. However, occasionally MDS will still occur even when phyllosomas are fed on enriched pipis. Obviously, some other factors, such as high density (less individual feeding) and inconsistent environmental conditions (inconsistent nutritional uptake) are inter-related in some way with MDS.
Diseases originating from environmental (physical and chemical) stress are not contagious. Environmental stress can manifest as white spots, with particles in the hemolymph clumping together. A high level of chemical contamination (chlorine, formalin etc) and physical stress (high stock density, freshwater drops from the cover) is the likely cause. Phyllosomas will die within 24 hours of the appearance of these white spots. There is no treatment.
POST LARVAL (NISTO) REARING
After metamorphosis to the nisto stage, nistos are reared in a nisto tank. The exoskeleton of nistos is transparent and not calcified. The colour of nistos changes from transparent-white to orange in colour due to the development of pigmentation under the exoskeleton. Water quality in the nisto rearing tank is the same as that of phyllosoma rearing. Nistos can be reared at a high density ( greater than 100 nistos per l). During the nisto stage, feeding is not required. The duration of the nisto stage is approximately 7 days, with the temperature kept at 26-27xc2x0 C. To avoid cannibalism, separate pre-metamorphosis phyllosomas from other intermoult phyllosomas. The same design as the larval rearing tank may be used for nistos, such as a one-way circular tank. The water should be treated in the metamorphosis tank in the same manner as the larval rearing tank. No feeding is required. No aeration is required.
JUVENILE CULTURE
After 7 days, nistos moult to the juvenile stage. Moulting to juveniles always occurs during the night. The exoskeletons of juveniles are calcified and pigmented. Newly moulted juveniles should be collected from the nisto tank the next morning and transferred to the juvenile holding tank.
Juveniles are nocturnal. Preferably, feed only once a day in the evening and clean the remaining food and faeces out the next morning. Adjust feeding levels according to the amount of remaining food. The food size for first instar juveniles is similar size to that of second or third instar phyllosomas. Chopped flesh of enriched pipis is suitable for up to at least fourth instar juveniles, then non-enriched pipis, squids, scallops and mussels can be used. The optimal temperature for culturing juveniles is 26-27xc2x0 C.
BROODSTOCK
After catching, live berried females may be stored in a tank within the vessel, preferably equipped with gentle aeration and/or water exchange. Feeding is not required. Environmental conditions, particularly water temperature and salinity, should be kept constant during the storage of the live animals.
Berried females should then be transported with water. Although berried females can survive without water for around 30 hours, long-term air exposure is physically stressful and occasionally berried females will scrape the eggs from their abdomen several days after delivery to the hatchery.
Small quantities of berried females ( less than 6) can be transported in a plastic bag packed with water (10-20 l) plus pure oxygen. The plastic bag is then placed in an esky, and sent to the hatchery. Berried females and their eggs can travel between 24-36 hours in the bags without ill effect.
For a larger quantity of berried females ( greater than 6), a fish transporter is recommended. More than 1 l of seawater per animal is an appropriate volume. During transport, pure air or oxygen should be supplied at more than 2 l per minute. Up to 24 hours of travel by this method will not adversely affect the berried females or the eggs.
The berried females are preferably kept in a holding tank with a seawater exchange of more than 100% per hour with aeration ( greater than 2 l air per min). A minimum of 30 l of seawater per animal is required. The seawater is preferably sterilized by UV before use. The tank is preferably covered such as with a black plastic sheet. The temperature of the water in the holding tank is preferably kept between 20-28xc2x0 C., with it being particularly preferred to maintain 26-27xc2x0 C. Daily temperature variation should no be greater than 1xc2x0 C.
The females may be fed once a day in the evening, and preferably with cleaning the remaining food out the next morning. Preferably, the food is selected from the flesh of molluscs, such as pipis, squids and mussels. The food is preferably sterilised by 0.1% chlorine solution for at least half an hour, and then carefully washed by UV sterilised seawater before feeding.
BROODSTOCK HOLDING SYSTEM
The broodstock holding system may comprise a round or square tank with a capacity of more than 200 l. There is preferably provided a black cover sheet on the top of the holding tank to minimise stress of broodstock. Gentle aeration may be provided, typically at about  greater than 2 l air per minute. Seawater may be supplied from one end of the tank, preferably from the bottom of tank, and discharged from the other end, preferably from the top of tank.
The water supply is preferably 1-5 xcexcm filtered and UV sterilised water, with a minimum of 30 l seawater per animal. An exchange ratio of  greater than 100% per hour is preferred, at a temperature between 20-28xc2x0 C. and preferably 26-27xc2x0 C.
Females are preferably fed once daily in the evening with food clean out the next morning by such as by siphoning, and scraping the bottom with a sponge. It is preferred to change the tank to new clean tank every 10-15 days.
HATCHING SYSTEM
The hatching system may comprise a round or square tank, typically having a capacity of 100-200 l. Seawater may be supplied from one end of the tank, preferably from bottom. Water may be discharged through a 500 xcexcm mesh to prevent escape of larvae. Preferably, there may be a black cover sheet on the top of the holding tank, with a 15-20 cm opening. It may be desirable to provide slow aeration of approximately 2 l per minute around the outlet. The water supply should be 0.5-1 xcexcm filtered and UV sterilised water. Preferably, an exchange ratio of approximately 100% per hour is used, with temperature preferably kept at 26-27xc2x0 C.
The hatching tank should be prepared in the afternoon, and the females transferred in the late afternoon. Feeding is not required in the hatching tank. The hatching tank should be sterilised by 0.1% chlorine for a period of 6 hours after the harvesting of larvae. If larvae have not hatched out by the next morning, females should be returned to the holding tank, and another hatching tank should be set up.
When the embryos become visibly amber-brown in colour, the individual berried female may be transferred to a 100-200 l hatching tank. The hatching tank should be prepared in the afternoon, and the females transferred in the late afternoon, prior to larvae hatching. Seawater may be supplied, previously filtered and sterilised by UV, at a rate of approximately 100% exchange per hour, exiting from a tank through a 500 xcexcm mesh to prevent escape of larvae. Preferably, slow aeration is supplied at approximately 2 l per minute around the outlet.
Hatching always occurs at around sunrise. When the eggs hatch, the female flicks her tail several times, and larvae are scattered into the water. This lasts about 10 to 20 minutes. Hatching occurs only in the morning, and occasionally spreads over two to three mornings. To minimise stress of the female, lighting should be avoided.
After hatching, larvae are very soft and fragile, and lack swimming ability and therefore strong aeration should be avoided. The exoskeletons of the larvae become hard, and they start swimming toward a light source 20-30 mins after hatching. Harvesting of larvae is possible when they congregate at the surface of the water (light source).
Larvae can only be transferred with water. A glass beaker or glass bowl is an appropriate vessel for harvesting.